Although most corn fields have been harvested, fields still standing should be checked for lodging potential. The dry weather experienced by many areas during the summer and autumn created stress on the plant, which can make it more susceptible to stalk rots and root rots during the grain fill period. There have been some reports of stalk rots and rotted roots, creating situations where a strong wind can knock the field over.
Lodging makes harvest more difficult, and it results in losses of grain since it cannot all be harvested. Furthermore, ears in contact with the ground are more likely to develop ear molds, reducing the quality of the grain and increasing the risk of contamination by mycotoxins. Mycotoxins are toxic substances produced by fungi.
A simple way to scout for lodging potential is to walk through the field and push the stalks 8-12 inches from vertical at chest height. Those stalks that fail to spring back can potentially lodge in a strong gust. If 10-15% of the field shows lodging potential, it is advisable to harvest that field as soon as is feasible.
For fields that exhibit lodging problems this year, it is a good idea to review plant populations and fertility programs to be sure they are not excessive.
For information about corn pests, visit
"Insect Management Recommendations".
Whiteflies in the greenhouse can be one of the more difficult to control insect problems to manage. In
Kentucky greenhouses, we have seen several species of whiteflies including greenhouse whitefly
(Trialeurodes vaporariorum), bandedwing whitefly (Trialeurodes abutilonea), and sweet potato
whitefly (Bemisia tabaci). We may also have the silverleaf whitefly (Bemisia argentifolii). Some
greenhouse plants are much more suitable for whiteflies than others with fuchsias, poinsettias,
ucumbers, lettuce and tomatoes preferred by whiteflies. Through regular monitoring, these preferred
hosts can be used as indicator plants, alerting greenhouse managers to the first signs of whitefly
nfestations.
These powdery white insects, about 1/12 inch in length, flutter from the undersides of leaves when the plants are disturbed. The lower surface of the leaves may be infested with all life stages of whiteflies. The female of these sap-sucking insects may lay 150 eggs at the rate of 25 per day. The newly emerged crawler moves only a short distance before settling down to feed. After three larval molts, the pupal stage is formed, from which the adult emerges. The entire life cycle takes 21-36 days, depending on the greenhouse environment.
The greenhouse, sweet potato, and silverleaf whiteflies are similar in appearance but differ in their biology and control. Whiteflies species develop entirely on the undersides of leaves. Their life cycle may be as short as 20 to 25 days. The sweet potato whitefly has a broader host range, higher reproductive potential, stronger resistance to insecticides and a powerfully phytotoxic enzyme system. This whitefly is a vector of gemini viruses in tomatoes. Control of these viruses relies on proper cultural controls and control of the whitefly vectors.
Identification of the adult stage is difficult, but there is a small difference in body color, the greenhouse whitefly having a body not as yellow as the other two. The greenhouse whitefly also holds it wings more horizontally at rest than the sloped arrangement of the sweetpotato and silverleaf whiteflies. The last instar nymph and pupa is the best stage to use for identification, these are usually found on the undersides of the older leaves.
Management of whiteflies begins with good cultural practices in the greenhouse. Solely relying on insecticides for whitefly control will not be effective.
Cultural Practices
Exclusion: We need to exclude whiteflies from the greenhouse as much as practical. Ventilators should be screened and doors closed when not in use. This is your first line of defense.
Isolation: When bringing in new plants from the field or plugs or plants that have been purchased from another greenhouse, isolate these plants in a separate section of the greenhouse until you determine they are not harboring unwanted insects or disease. Yellow sticky cards hung just above the height of the plants works well as a monitoring tool. Yellow sticky cards will attack adult whiteflies, thrips, aphids, and fungus gnats.
Sanitation: Keep the greenhouse free of weeds that may harbor insects and diseases from one cropping cycle to another. The area around the outside of the greenhouse needs to be keep weed free as well, particularly around the doors and ventilators. Try to avoid keeping 'pet' perennial plants in the greenhouse. These may harbor insects and diseases without showing signs of a problem.
Monitor Regularly: Weekly inspection of the plants is required to stop small problems from becoming large problems. Yellow sticky cards near doors and windows can provide an early heads-up to potential problems. Plants that are light to moderately infested should be isolated from uninfested parts of the greenhouse. Heavily infested plants should be carefully discarded. When monitoring, be sure to thoroughly examine plants, as many of the greenhouse pests occur under the leaves, in the buds, or in growing tips of the plants. A hand lens may be necessary. Always visit the suspected 'hot spots' last, as it is easy to carry insect and mite pests on your clothing to uninfested parts of the greenhouse. Avoid wearing bright colored clothing, as some insect pests are attracted to bright colors and can be carried into the greenhouse. Certain plants are very susceptible to specific insect and mite problems, those plants can be used as indicators of developing problems.
Biological Control: Greenhouse operators should consider biological control for recurring problems. Encarsia formosa remains one of the more effective controls for greenhouse whitefly, although it is not as effective for sweet potato or silverleaf whitefly. Eretmocerus eremicus, another parasitic wasp, and Delphastus pusillus, a small predatory beetle are other biocontrol agents that have been evaluated for whitefly control in the greenhouse.
Plant Spacing: Often it is necessary to use insecticides for whitefly control. Non-systemic insecticides require complete coverage of the plant, so plant need to be spaced accordingly.
Insecticidal Control
Know Your Insecticide: Different whitefly insecticides will work differently. Some are effective against the eggs, nymphs and adults, others may target the adult stage. There are several insect growth regulators (IGR's) for whitefly control, these are effective against the immature stages, eggs, nymphs, and pupae. The IGR are often used early in the cropping cycle to delay/prevent whitefly development.
Coverage: As the immature stages of whiteflies occur on the undersides of leaves, sprays of non-systemic insecticides need to cover both the upper and lower surfaces of the leaves. Immature stages are not mobile and will not move to contact insecticide residues.
Resistance: Insecticide resistance remains a very critical issue with whiteflies. Sweetpotato and silverleaf whiteflies have already developed resistance to many classes of insecticides. When insecticides sprays are used to control whiteflies, classes with different modes of action should be rotated every 18 to 28 days. As coverage is difficult, it is often necessary to make multiple applications of non-systemic insecticides at 3 to 7 day intervals to achieve control. Do not tank mix insecticides with the same mode of action, as this is of little value. Many of the newer insecticide labels now indicate the chemical group an insecticide belongs to.
Unintended Effects: Some insecticidal sprays are not compatible with biological control agents. Prior to release of natural enemies in the greenhouse, insecticidal soap can be used to reduce pest numbers. Some plants or plant stages may be sensitive to certain insecticides, so select insecticides accordingly and read the label.
In a previous article, we discussed the role of sanitation and eradication in the management of diseases in the greenhouse. This week, we'll cover the principle of exclusion and the role it plays in disease control. The recent discovery of an unidentified, whitefly-transmitted Begomovirus (geminivirus) in greenhouse-grown tomatoes in Louisville, the first known occurrence of this plant virus in Kentucky, is a potent reminder of what can happen when a pathogen is introduced in the greenhouse environment.
One of the biggest threats to agricultural production around the world is the introduction of pests through global movement of plants and plant products. The goal of exclusion is to prevent a pathogen from being introduced into an area where the pathogen does not already exist. This can be accomplished through quarantines or by the production and use of pathogen-free seed and transplants. When plant material is brought into a greenhouse from an outside source, don't assume that this material is pathogen- (or insect) free. Keep imported plants in a separate greenhouse for 2-3 weeks and monitor them for symptoms of disease or emergence of insect vectors such as whiteflies and aphids. Regular scouting is required, and sticky cards can also be employed to monitor insect populations. By keeping imported materials in isolation, the likelihood of a disease spreading into production areas is lessened. Should a disease be suspected on plants in quarantine, contact the Plant Disease Diagnostic Lab in Lexington or Princeton, or call your extension plant pathologist. When sending samples into the clinic, double-bag them to prevent release into the environment. Certain pathogens could threaten crops in the field if released, depending upon the time of year, and could be devastating should the pathogen become established. Quarantined plants that develop disease should be destroyed promptly, and every attempt should be made to eradicate insects that are carried on these plants. More information on insect control can be obtained from the Dept. of Entomology at the University of Kentucky.
Prevent movement into the greenhouse of infested soil on equipment, implements, shoes, and clothing. Wash hands and sanitize shoes when going from greenhouse to greenhouse. Destroy or sanitize tools and materials, etc. that have come into contact with plant pathogens. Don't use surface water to irrigate in the greenhouse, as this can introduce pathogens into the system. Do not grow vegetable transplants in facilities where ornamental plants are produced, as ornamentals may harbor pathogens (especially viruses) and their insect vectors that can then spread to vegetable transplants.
The very nature of today's global economy provides the ideal set of circumstance for non-native and potentially destructive pests to be introduced into the U.S. Protect your operation and others in Kentucky and the nation by including the principle of exclusion in your pest management plan.
Phytophthora ramorum is a fungus-like organism that can infect woody trees and shrubs, herbaceous plants and ferns, causing diseases like Ramorum leaf blight and Ramorum shoot dieback. On mature oak and tanoak trees this pathogen causes sudden oak death. This organism was found in the United States initially in forested areas in coastal California in the mid 1990's. Later it was found to be present in a forest area in Oregon and in nurseries in California, Oregon and Washington. It has also been found in nurseries in several countries in Europe and in British Columbia, Canada.
On plants like rhododendron, camellia and viburnum leaf and twig symptoms like brown lesions and blotches on the leaves and necrosis of the leaf tips and leaf blight are typical symptoms of Ramorum blight. Shoot necrosis and dieback are typical of Ramorum shoot dieback and are seen on rhododendron, blueberry and on conifers. Some plants like camellias shed the infected leaves, and the disease is not immediately lethal to them. However, a large number of spores are produced on the leaves and the spores can infect oak and tanoak trees causing sudden oak death, a disease that can kill the trees in a relatively short period of time.
An updated list of host plants and plants associated with Phytophthora ramorum is available on the USDA-APHIS-PPQ website. The last revision was dated September 14th 2005, when eight plants were added to the list of P. ramorum hosts and plants associated with P. ramorum bringing the total number of plants listed to 85. Proven hosts are regulated in whole or in part and forty plants are listed as such. The other 45 plants are associated with P. ramorum and regulated as nursery stock. The complete list can be viewed at: http://www.aphis.usda.gov/ppq/ispm/pramorum/pdf files/usdaprlist.pdf.
The long distance spread of Phytophthora ramorum is believed to occur through the movement of infected plants, plant parts and even soil. There is great concern that P. ramorum may spread to parks and native woodlands in the U.S. from introduced infected ornamental plants and even soil, and that native plants in forest areas outside the areas presently known to have the pathogen and sudden oak death may become affected. Nursery and forest surveys are under way in the United States, Canada and Europe for detection of this pathogen.
In the United States, national nursery surveys have been completed in 48 states. Seven states were found to have nurseries with plants that tested positive for P. ramorum and the number of nurseries in each of the seven states were: 55 in California, 20 in Oregon, 13 in Washington, four in Georgia, two in Louisiana, one in South Carolina, and one in Tennessee. In 2004 infected plants were found in nurseries in 22 states. These numbers show a reduction in the number of nurseries with infected plants outside of the three regulated west Coast states, California, Oregon and Washington. Fortunately however, this is not the case for the regulated states, where the number of nurseries with infected plants has increased.
In the state of Kentucky a nursery survey was performed during the summer of 2005, as a collaboration between the USDA-APHIS, the Office of the State Entomologist and the Department of Plant Pathology at the University of Kentucky. Nurseries were inspected by USDA-APHIS personnel and nursery inspectors operating from the Office of the State Entomologist, and samples from plants showing symptoms similar to those expressed by plants infected by P. ramorum were collected. Samples were placed in double bags with zip closure, labeled and sent to the Plant Pathology Department of the University of Kentucky for analysis. All samples were tested for infection by Phytophthora using an enzyme linked immunosorbent assay (ELISA). This assay uses antibodies that recognize proteins present in organisms in the genus Phytophthora but do not recognize species. There are approximately 50 species in the genus Phytophthora and any samples that were ELISA-test positive were tested further. Total DNA was extracted from the suspect positive sample and the DNA was sent to the USDA-APHIS for testing by polymerase chain reaction (PCR) using nested primers for P. ramorum DNA amplification. This PCR technique is at present one of the two best methods for identifying P. ramorum.
Approximately 30,217 plants in 105 nurseries and retail outlets were surveyed in 33 counties. There were six trace forward nurseries in Kentucky, that is, nurseries that received camellias from a contaminated west coast nursery in 2004, and these were also surveyed. A total of 26 samples were collected from nurseries in the following ten counties: Boone (5), Campbell (4), Clark (1), Fayette (3), Hardin (1), Jefferson (7), Jessamine (2), Madison (1), Pulaski (1), Taylor (1). Two samples were positive in ELISA for the genus Phytophthora. DNA was extracted from these two samples and was sent to USDA-APHIS lab for PCR and the result was negative for P. ramorum.
One rhododendron plant showing symptoms similar to those caused by P. ramorum was found in the survey of the six trace forward nurseries. A sample was collected from this plant and it was negative in ELISA for P. ramorum.
No samples collected from nurseries in the state of Kentucky were found to be positive for P. ramorum in 2005. A forest and nursery perimeter survey was also performed in Kentucky in 2005, and no suspect samples collected were found to be positive for P. ramorum. Early detection and eradication of diseased plants are important to protect Kentucky's forest resources and the landscape and nursery industries, and surveys like the nursery survey and the forest survey are important in the effort to achieve this goal.
Root and butt rot fungi affect many kinds of landscape trees in Kentucky, causing decays of the roots and lower trunk and leading to dieback and decline. Two diseases that have been observed this summer and fall include decays caused by Ganoderma and by Xylaria. The fungus Armillaria which also causes root and butt rot, especially in trees stressed by drought or insect defoliation, was discussed recently in this newsletter (KPN # 1073, dated August 8, 2005). These three diseases are readily diagnosed by the kind of fungal fruiting structure produced at the base of the trunk or from the tree's infected roots. There are no cures for any of the root and butt rot diseases of landscape trees.
Ganoderma root and butt rot caused by the fungus Ganoderma lucidum, incites loss of vigor and undersized leaves or dead branches in infected trees. Basidiocarps (fruiting structures) of this fungus are annual, a few inches to a foot across, and leathery to corky when fresh. The upper surface has a smooth, hard, varnish-like coating with a red-brown color. The margin and under surface is white or off-white and contains pores. In the landscape, basidiocarps are sometimes obscured by turfgrass, mulch, or ground cover plants growing at the base or over the roots of the affected tree.
The fungus causes a light-colored, spongy sapwood decay which occurs gradually in the tree roots and butt, and affects their ability to function in transport of water and mineral elements in the tree, leading to decline. In addition, the decay affects the strength of the tree making them subject to falling in stormy weather. In the landscape, trees can sometimes appear to have healthy foliage and branches while harboring significant root and trunk decay. Despite their appearance, such trees may pose a risk to people and property and need to be evaluated by an ISA-certified arborist to determine the extent of decay and to help the tree owner with decisions about tree removal.
This disease is commonly seen in Kentucky on apple, honey locust, and oaks. In addition, ash, cherry, elm, hickory, hornbeam, Kentucky coffee tree, magnolia, maple, redbud, sassafras, and sweetgum are susceptible to Ganoderma rot.
Xylaria root rot is also called black root rot (not to be confused with black root rot caused by the fungus Thielaviopsis basicola) because of the black sheath of fungal stroma found on the surface of decaying roots. Two species of Xylaria, X. mali, and X. polymorpha are the causes of this root rot disease. These fungi produce an off-white decay of tree roots, leading to the decline and loss of strength typical of root and butt rot diseases.
Xylaria produces clusters of black, finger-like or club-shaped fruiting structures that originate on the lower trunk or on major roots. These fungal structures, a few inches long, resemble blackened "fingers," inspiring the common name, "dead man's fingers." These structures are unique and easy to recognize when they appear at the base of the tree or in the lawn or landscape bed nearby.
This disease is seen in Kentucky on apples, honey locusts and yellowwood. Additional hosts include cherry, elm, maple, and possibly beech, hickory, black locust, oak, sassafras, and walnut.
Twig girdlers are species of longhorned beetles that have a very distinctive approach to laying their eggs. Females select twigs about the diameter of a fat pencil and chew deep, narrow grooves that leave about a 2 foot-long section attached by only a slender piece of heartwood. The brown beetles crawl along the terminal portion and make small notches in which the eggs (about 5 to 20) are placed. Girdled twigs, which contain eggs and white legless larvae, soon break and fall to the ground. The results of this handiwork can be seen littering the ground under a variety of trees including hickory, pecan, and oak. Twig pruning produces growth deformities that affect the shape and appearance of small trees.
Collection and destruction of fallen twigs is the most effective means of reducing the potential infestation for next year. Application of insecticides to control these insects has not been very satisfactory.
Recent samples in the Diagnostic Laboratory have included Penicillium ear / kernel rot and Fusarium stalk rot on corn; and downy mildew and sudden death syndrome on soybean.
On fruit and vegetable samples, we have diagnosed cork spot associated with calcium deficiency and sooty blotch/flyspeck on apple; downy mildew on cucumber; Fusarium fruit decay on pumpkin; and Septoria leaf spot and yellow shoulders (physiological disorder) on tomato.
On ornamentals and turf, we have seen foliar nematodes on anemone; Rhizoctonia web blight on chrysanthemum; Botryosphaeria canker on ash; black root rot on holly; bacterial leaf scorch on oak and sycamore; black spot on rose; Rhizosphaera needle cast on blue spruce; gray leaf spot on fescue; and Ustilago smut on crabgrass and fescue.
View Princeton trap counts for the entire 2005 season at - http://www.uky.edu/Ag/IPMPrinceton/Counts/2005trapsfp.htm
Fulton County trap counts are available at -http://ces.ca.uky.edu/fulton/anr/Insect%20Counts.htm
For information on trap counts in southern Illinois visit the Hines Report at - http://www.ipm.uiuc.edu/pubs/hines_report/comments.html The Hines Report is posted weekly by Ron Hines, Senior Research Specialist, at the University of Illinois Dixon Springs Agricultural Center.
NOTE: Trade names are used to simplify the information presented in this newsletter. No endorsement by the Cooperative Extension Service is intended, nor is criticism implied of similar products that are not named.
Lee Townsend
Extension Entomologist